Lab safety

You must always wear long pants, closed-toed shoes, and a lab coat when in the lab!

Always use gloves when working in the lab but DO NOT WEAR GLOVES OUTSIDE OF THE LAB (e.g. hallway).

Pre-PCR and post-PCR are carried out in specified areas in the lab, Moffett 333A. Discard gloves when moving between areas and keep all equipment in their designated rooms. AVOID moving things back into the pre-PCR area. These precautions are to avoid contamination.

If you use the last of the supplies (such as tips or gloves), please restock these items.

When stock solutions, kits, gloves, Kimwipes, etc runs out, it is your responsibility to make sure these get replaced. If more needs to be ordered, please notify the appropriate person.

At the end of the day, remove all of your stuff from the bench you were using and return all equipment to their appropriate places, washing glassware with deionized water and hanging on the rack to dry.

Please turn off equipment if it is not in use. Especially hot plates! Make sure they are off when you are done using them.

If you uncomfortable at all with any laboratory technique, please do not attempt the procedure and ask for guidance.

Sean Grace in EEB can help with needs relating to research and facilities! Make friends with him! He likes cookies.

IThelp@princeton.edu will help with all things computer and software-based. I can purchase software if you need but printer drivers, computer beachballs of death, etc.. they are your friends.

Ice is found is Moffet 425

Your supervisors (graduate student or post-doctoral researcher and Dr. vonHoldt) are responsible for you. Please make every effort to be a good laboratory citizen. The vonHoldt lab is filled with friendly and knowledgeable people. Please ask questions when you are at all uncertain about your technique, procedure, or project. Please make every effort to attend our laboratory meetings, as this is an important forum for meeting the lab members and keeping up-to-date on issues in the lab (i.e. new techniques, current research projects underway, presenting your own work, etc.).

Lab Safety Training
Please log on to Princeton's Environmental Health and Safety webpage to learn about lab safety and other protocols. A University laboratory safety manual is available online for your viewing. The vonHoldt lab, however, has some specific protocols and training that needs to be recorded for each individual in the lab to ensure your safety while working at the lab bench. In addition, we also have a Departmental EEB Chemical Hygiene Plan. Each person needs to complete two training sessions

Laboratory Safety (3 hour session - go to the LearnCenter for enrolling in the course)

5M Sodium Chloride (NaCl) (makes 250mL)
1. Obtain a beaker with at least 250mL capacity
2. Measure out 73.05g of solid sodium chloride (NaCl) and add to beaker
3. Add distilled water up to total volume of slightly less than 250mL
4. Place beaker on stirring block, add stir bar
5. Stir and heat slightly
6. Add water until total volume is 250mL and sodium chloride is totally dissolved
7. Remove stir bar with magnet and place solution in a 500mL bottle with a loosely fitted cap
8. Autoclave (liquid cycle: ~20 mins)
9. Once cooled, run solution through sterile filter system

Obtain a short length of plastic hose to connect vacuum line to filter unit

Make sure receiving unit (bottom bottle) and filter unit (top half) are held together tightly

Qubit quantification of double-stranded DNA

Qubit gives a more accurate reading of the amount of dsDNA in a sample. Because we are using a High Sensitivity (HS) kit, your samples should be 1-100 ng/uL to ensure accuracy. You can use anywhere from 1-10uL of sample in each tube, always with a total volume of 200μL. We use 2μL to save sample and avoid potential pipetting errors of using 1μL.

The dye used in Qubit is light sensitive. Keep dye tube covered in aluminum foil. Put sample tubes with dye in tube rack covered with aluminum foil or in a draw when possible.

Materials

Qubit dsDNA HS Assay Kit

HS buffer

HS standard #2 (10ng/μL or 100ng/μL)

Quant-it dsDNA reagent (light sensitive dye)

Qubit Assay tubes

Tin foil

Methods
1. Thaw High Sensitivity buffer, light-sensitive dye, standard and samples to room temperature (or use room temperature stock).
2. Cover a tube rack in aluminum foil for storing samples (or place in a drawer).
3. Pull Qubit tubes for number of samples plus two for the standards.
4. Label tubes STD-1 and STD-2, and label one tube per sample you will quantify. Put in tube rack covered in aluminum foil (or drawer).
5. In a sterile 1.5 mL tube covered in tinfoil mix the following. You should master mix this for the number of samples plus three to include standards and extra.

Nanodrop 2000 quantification of DNA

Please use your own materials (pipette, pipette tips, Kimwipes, etc). Start NanoDrop 2000 software and select analysis method (e.g. Nucleic Acid for DNA). Carefully apply a droplet of water (1-2µL) to the pedestal to clean and initialize the instrument. Use a Kimwipe to clean the pedestal and the top in between readings. At the prompt, name the results file and save it in the “My Documents” folder. Apply 1-2µL to the pedestal and press the “Blank” button (AE buffer, TE buffer, water, etc). Clean the pedestal and apply 1-2µL of sample, then press the “Measure” button. After measuring, the concentration and other data appear in the software window. Continue measuring remaining samples. After measuring, view the results table.

Polymerase Chain Reaction (PCR): General Introduction

A PCR contains the following necessary reagents:

PCR-buffer. Salt and pH-stabiliser. User stock of 10x is kept in your box.

MgCl2. Salt which is required for the Taq polymerase to work. The standard rxn concentration is 1.5 mM (range 1-4). Higher concentrations makes the Taq polymerase less specific and favours amplifications of short fragments. Too much MgCl2 often results in multiple bands. User stock of 25mM is kept in you.

dNTPs. Free nucleotides (Gs, As, Ts and Cs) of which the artificial DNA copies are made. User stock (10 mM of dNTP Mix, which has 2.5mM of each dNTP) is kept in your box.

Primers. Single stranded DNA (oligonucleotides), usually the length of 18-30 bp. Primers used for RAPD are normally shorter, 10-15 bp. Stock solutions at 100µM are normally, and user stocks at 10µM are stored in your PCR box.

Taq DNA polymerase. The enzyme that puts the free nucleotides together. It starts at the 3' end of the primer and uses the complementary DNA strand as a template. User stock of 5 units/µl is kept in your box.

Template DNA. The source of DNA for the PCR amplification. This could be DNA extracted from blood, skin, feathers, or old PCR products. We use a standard concentration at 25 ng/µl but depending on the organism and protocol, the concentration might need further adjustments (5-100 ng/µl).

In addition to these reagents, you may find that researchers are adding other reagents to their reactions in hope of getting better results. You may consider:

To set up a PCR:
1. Make sure you have reserved a thermocycler for your usage. The sign-up dry-erase board is in 5202.
2. Create a PCR protocol and calculate how much you need of your master mix/ cocktail (number of rxn + 10%). Don't forget the rxn for the blank (negative control). The master mix should contain everything except the template (but you should add the Taq just before aliquoting).
3. Make sure the PCR bench is clean.
4. Thaw template DNA and all reaction reagents on the PCR bench, except the Taq polymerase which should remain in the freezer until needed (it contains glycerol so it does not need thawing).
5. Program the PCR machine while the reagents are thawing.
6. Mix and spin all reagents and keep on ice (once experienced with the technique, keeping all reagents on ice is not necessary, unless there are unexpected delays).
7. Place an appropriate number of PCR tubes in a tray, and briefly label each row of tubes.
8. Aliquot your desired amount of template DNA prior to adding Taq polymerase to your cocktail.
9. Make the cocktail in an 1.5 ml eppendorff tube (snap cap). Start with adding the ddH20 and save the Taq DNA polymerase to the last step. Mix by using the 200 µl pipette set at 150µl and pipette up and down a few times.
10. Dispense an appropriate volume of master mix into each of the reaction tubes (total rxn minus amount of template DNA).
11. Add the template (change tip between samples!!!!).
12. Fix the lids on the tubes.
13. Bring the tubes over to the thermocyclers in room 4162 (on ice if you haven't programmed them yet--it may take a few minutes for a newly turned on machine to warm up). Start the PCR machine. Select and run your program, and ALWAYS USE A HEATED LID! Load the tubes, close and tighten the lid, then you are ready to go!

Programming the thermocycler (choosing a temperature profile):
The standard PCR starts with a warming up phase of 3 minutes at 94˚C, to make the template DNA single stranded (denatured). Then follows the cyclic phase that characteristically consists of three different steps.
1. 94˚C. This is again the denaturing step that initiates all cycles and is normally set between 30-60 sec.
2. 37-70˚C. The annealing temperature when the primer is allowed to settle on the template DNA. This step is usually set between 30-120sec. The chosen temperature depends on the melting temperature, Tm, of the primer (length and GC-content).
3. 72˚C. The elongation temperature is the optimal working temperature of the Taq DNA polymerase. This step is set between 5-500 sec depending on the length of the desired fragment. A rule of thumb is that the Taq polymerase builds about 1,000 nucleotides per minute.
4. The number of cycles used varies normally between 20-40 depending on the template DNA concentration, quality, length of product, and above all, empirical experience with the focal reaction.
5. The reaction is normally ended by a 10 minute phase at 72˚C. This will allow the Taq polymerase to add a protruding A at the 3' end of the fragments. This step is very important when cloning the PCR fragments by means of TA-cloning.

Recipe for 2% agarose gels (small gel chamber):
1. If a 2% solution is already made up, proceed to step 3 to melt the gel. To make 100 ml of new solution, take weighing paper from the drawer, place it on the balance, tare, and add 2.0 g agarose (on shelf above microwave). Add to flask.
2. Add 100 ml of 1x TAE buffer (in carboy).

200 ml 1x TAE with 4g agarose for large gel

3. Place the flsk in the microwave and boil for repeated 30 sec intervals until the agarose has melted (about 2 min). Make sure it doesn't boil over! Use the orange heat-protection gloves when handling the warm flask.
4. Tape the ends of the gel casts and insert combs. Use two combs if you don’t need to run the fragments out the full length of the gel--it saves time and materials.
5. When the temperature of the agarose solution reaches 60˚C (you should be able to hold the glass for 5 seconds without burning your hand, but make sure the gel hasn't started to polymerize yet), add 3 µl of SYBR Safe to the agarose solution.
6. Pour the solution into the cast so that the gel is about 2 mm thick. Cover with box and allow it to polymerize (about 15-30 minutes).When polymerized, remove the tape.

Running 2% agarose:
1. Move the gel (still in gel cast) into the gel chamber with wells near the black electrode and remove combs.
2. If necessary, top off the 1x TAE buffer to the fill line of the chamber. The buffer can be reused several times, but should be replaced every second week.
3. Take parafilm and pipet 1-2µl of loading dye corresponding to the number of your samples plus one for a DNA ladder (if needed) (may be useful to use Post-PCR multi-channel pipet).
4. Add 5-20 µl * of the final PCR product to each well (change tips between samples) (may be useful to use Post-PCR multi-channel pipet).
5. Load the first well in the gel with 5 µl of a DNA ladder, and then the samples (PCR rxn and dye). You can reuse tips here--just place the tip in the buffer in the chamber and pipette up and down to flush the residue.
6. Put the lid on the electrophoresis chamber.
7. Turn on the power supply, adjust the voltage (80-100V), and let the gel run for 50-60 minutes.
8. When finished, place the gel on the glass plate in the UV camera box. For our camera box, the UV light turns off when the door is open. However, be cautious about exposing your skin to UV rays for too long: severe burns may develop without protective clothing or eyewear.
9. When finished, carefully dispose of the gel in the regular waste bin. Wipe down the glass with kimwipe and dispose of them in the regular waste bin.

If you need to clean your PCR product by exo-sap or sending to Beckman, load less of your PCR product (5ul) or use a surface tension gel to load even less.

Recipe for 2% agarose gel (Surface Tension):
1. To make 30 ml of new solution, take weighing paper from the drawer, place it on the balance, tare, and add 0.6 g agarose (on shelf above microwave). Add to flask.
2. Add 30 ml of 1x TAE buffer (in carboy).

50 ml 1x TAE buffer with 1.0g agarose for large glass plate

3. Place the flask in the microwave and boil for repeated 30 sec intervals until the agarose has melted (make sure it doesn't boil over!). Use the orange heat-protection gloves when handling the warm flask.
4. When the temperature of the agarose solution reaches 60˚C (you should be able to hold the flask for 5 seconds without burning your hand, but make sure the gel hasn't started to polymerize yet), add 1.5 µl of SYBR Safe to the agarose solution.
5. Put binder clips on each side of a comb.
6. Pour the solution onto a clean small glass plate and place comb in after you pour the solution.
7. Cover with box and allow it to polymerize (about 10 minutes).When polymerized, remove the comb.
8. Load 4 µl ladder with 1.5 µl (if needed) and 3 µl PCR product with 1.5 µl dye.
9. Run gel at 100V for 30-35 min.

A general PCR set-up template for keeping notes on the experiment, samples, thermocycler, and master mix.

A Brief Intro to Primer Design

During PCR, primers are annealed to complementary regions of single stranded molecules (a result of the denaturing step in the PCR cycle program). The primer sequences are then extended by the DNA polymerase in the PCR cocktail. Both of these steps are temperature sensitive and testing (or knowing) the right temperatures for your primers are testable. Primer annealing is often around 50-60°C but can be identified through testing.

Good primer design is essential for successful reactions. Here are some important design considerations:

Primer Length: Optimal length of PCR primers is 18-22 bp is a general guideline. This length is long enough for adequate specificity and short enough for primers to bind easily to the template at the annealing temperature.

Primer Melting Temperature (Tm): The temperature at which one half of the DNA duplex will dissociate to become single stranded. Primers with Tm in the range of 52-58°C are ideal, with Tm's above 65°C likely to have secondary annealing. The GC content of the sequence gives a fair indication of the primer Tm.

Repeats and long runs of a single nucleotide in the primer sequence: Avoid them! But if you need, no more than 4 of any is suggested.

Design your primer using the program Primer3. Also, one other fantastic method for discovering primers is from genome sequence. You can easily use the program msatcommander for microsatellite primer design based on a series of parameters. I suggest developing primers for tetra-nucleotide repeats and using M13 tags for a 2-step PCR and dye-label. This is cost-effective and flexible. Also keep in mind if you want to multiplex the primers eventually - that way you can be choosy for product sizes and also try to prevent physical linkage when possible to retain as many loci as possible.

Primer testing 'in-silico' using the UCSC Genome Browser (select PCR from the home page) or use this UCSC PCR link!

What are M13 primers, anyways?

A process where we can save money by not having to buy primers that are already dye-labeled, we instead add a 16mer sequence tag (there are many sequences and ensure your M13 matches the dye label....we use the M13F –20 sequence 5'-GTA AAA CGA CGG CAA G-3' or some variant of) onto the 5’ end of one of the primers (called a M13-hybrid primer). After the first couple rounds of PCR cycles, the 16mer tag gets added onto your copied DNA product. We put a small enough amount of the hybrid primer in the PCR cocktail so that it all gets used up by 20-25 cycles. We also put a small amount of M13F-20 primer (16bp in length) in the mix, and this primer has been dye labeled. We then drop the annealing temperature by 5°C and this allows the much shorter dye labeled M13F-20 primer (16bp) to anneal and be added onto the copied DNA strands. We run another 20 cycles and you end up with dye labeled PCR product that is 16bp longer than than your original primer sized product (good to remember if you are comparing it to results using the regular primer).

This is much cheaper because we can buy the M13-20 dye labeled primer in bulk, we need to use so little, and we do not have to buy unique dye labeled primers for every primer set we wish to use (the M13-hybrid primers cost about $10-$12, instead of $70 - $120 for a dye labeled primer).

What if I am using Fluorescently dye-Labeled (DL) primers for microsatellite PCR? (AKA setting up a multiplex after screening)

PCR and pooling can be made more efficient with DL primers; that is, when you order a forward primer sequence with a fluorescent dye labeled already attached. In the M13 protocol, the dye is not attached to the forward primer; it is attached during the 2-step PCR thermocycling program. Once you have screened your primers for amplification success and variability, you may decide on pooling 4, 8, or >10 primer pairs to construct a multiplex. A multiplex is typically constructed by color, size, or both. If just by color, then amplicon sizes can be overlapping but each primer pair that is pooled into the PCR has to have a different fluorescent dye label attached to it for downstream differentiation. If you choose to multiplex by amplicon size, then amplicons MUST NOT BE OF THE SAME SIZE but you could use all the same dye label (e.g. 6FAM) in the PCR. If you want to multiplex together by both size and color, then you can use colors to differentiate amplicons of the same size, while amplicons of the same color must be of different sizes.

Ordering DL primers is typically done so they arrive at a standardized concentration (e.g. pMol) so diluting them is typically done by just adding 100ul of di-H20 to reach a final concentration of 100uM. For example, they may arrive dried at 10,000 pMol. To hydrate them to 100uM solution, simply add 100ul of di-H20 to the tubes, vortex or flick, and let sit at room temperature for 10-20min before use. Word of caution: always double check their concentration before adding di-H20!

To make primer mixes, add 2uL each primer (forward and reverse – both at 100uM) and then fill the volume up to 100uL with water.

3. Add 9.5μL of the HiDi/Liz mixture per well of a 96-well plate.
4. Make a 1/20 dilution of PCR product.

2μL PCR product to 38ul water.

Use a cheap dilution plate.

5. Add 2μL of diluted PCR to the 9.5ul HiDi/Liz in each well.
6. Use a sticky lid and centrifuge.
7. Denature at 95°C for 5min (disable heated lid); "denatureMSATS" program.

Place on ice immediately for 5min.

8. Spin down plate and place back on ice.
9. Label plate with name: vonHoldt_<PlateName>, your initials, date.
10. Be sure to record the number of plates shipped to Cornell for budget purposes!
11. Use Geneious to genotype the microsatellites.

Microsatellite genotyping using GeneMapper v3.0/3.7

2. Set up marker panel and create a Bin Set in Panel Manager; set up bins if alleles are known. This is done in a hierarchical method in the Panel Manager:

In the left navigation window, click Panel Manager and then “New Kit” (upper far-left button). Name it and select the type of data you want.

Select on the new Kit you created, then click “New Panel”. Name it and press Enter.

Click on the panel you created, then click “New Marker”. Name it, provide marker range, color, and number of repeats, and comments if you choose.

Add in as many markers you have. I create different Kits for each multiplex combination set. Make sure to name everything consistently so you can link up to them later and you know exactly what it means. Example: name Analysis Method, Panel, Kit, etc all matching the microsatellite multiplex primer mix so you can easily cross-reference them when setting up in GeneMapper.

Next, select your Kit and in the Bins menu, select New Bin Set. Name it and then you can select it from the drop-down menu. This Bin Set is used in the Analysis Method to again, be clear about how you name your Kits and Panels.

3. Set up Analysis Method specific to your samples in GeneMapper Manager.

In the GeneMapper Manager, click the Analysis Method Tab, and then click New.

Follow prompts, then name your Method on “General” tab.

“Allele” tab: this is where you link your Analysis Method to the Bin Set you just created. You can also click “Use the marker-specific stutter ratio” if your marker stutters.

If marker is known to stutter, you can the change the stutter ratio (value is a percentage of the time you observe stutter in dataset) and this will avoid “over labeling” of stutter peaks.

“Peak Detector” tab: click “User Specific (rfu)” and here you can set a Minimum Peak Height requirement; useful to avoid over labeling low-intensity peaks. The higher the value, the higher the intensity requirement for peak calling.

The remaining tabs have tons of values you can change to better tweak your analysis. I haven’t figured them all out yet, but don’t let that stop you from having fun!

Click OK, then Done.

4. Select your Analysis Method, Panel, and Size Standard information on Samples Tab of project, and hold “Ctrl-D” to fill down for all samples.

6. On the Genotype Tabs, each individual is represented by a separate entry for each marker it was run with (if you multiplexed markers). Each run is tested for quality control in many categories. The GQ (Genotype Quality) column is most important and is rated one of three quality levels:

Green square: good to go!

Yellow triangle: usually ok to go!

Red hexagon: bad; it’s a no-go!

7. To view all electropheregrams, select all runs in each marker (left navigation window in Genotypes Tab) and click the “Display Plots” button.

8. There are two important buttons in the upper left corner (Peak Selection Mode and Binning Mode buttons). You can view runs and add allele bins here if necessary.

Peak Selection Mode: Here you can click on any peak, add it as an allele you created previous to the analysis run, delete the allele call, or custom name the allele peak as a “new” allele for that individual peak.

Binning Mode: Here you can create/delete/edit bins and increase/decrease the marker range as needed. Any changes made are saved when you exit the screen but you will need to RE-analyze for it to be applied to all the other samples analyzed with that marker. Go back to the Samples Tab and click the “Analyze” button.

9. I recommend viewing all runs to double check the calls made by GeneMapper. If you think a run is good, but the program gave it a GQ red hexagon (failed), you can override this by two ways:

When you make any changes to a run, it automatically replaces quality values with gray triangles. This tells you that you made manual changes to the run.

You can “right” click on the final GQ value and a prompt appears asking “Override the Genotype Quality of this marker?” Yes will also create the gray triangles.

10. Once you have viewed/edited all the runs for a single marker, you can always sort the entries under the Edit menu. You can also edit the Table Settings in Table Settings Editor to hide some of the excessive columns that normally appear on both the Samples and Genotype Tabs. Make sure to select the Table Settings you created in the drop- down menu in order for it to be applied to your project.

11. To Export your Allele Table, click “Export Table” button and save it in your folder. It is saved as a “.txt” file and easily copy/pastes into Excel. No problem!

12. Congratulations! (as a last note, I recommend re-analyzing all samples for a marker when you add more samples for that marker...keeping allele calls consistent and standardized across PCRs).

Troubleshoot:

Check that you are viewing the table in the correct table edit selection. Microsatellite data should be under the Microsatellite Default viewing screen, AFLP with the AFLP screen, and so on.

Check that you have selected the appropriate bin set when viewing alleles. This makes a world of difference.

Design as small of amplicons as possible (target product size of 45-80 bases max)

The SHORTER THE AMPLICON THE BETTER

Keep decreasing product size until Primer3 can’t design primers

Name the primer set: HRM-chr#.position (example: HRM-chr15.32383555)

Setting up the HRM plate and master mix

Reagents for the HRM Mastermix cocktail (1x)

1x

Primer mix (2μM or 4μM - this is up for discussion)

0.5μL

Roche MgCl2

1.75μL

diH2O

2.25μL

Roche Master Mix (pre-made from the kit and light sensitive!)

5μL

DNA 1ng+

1μL

Total

10.5μL

Since we are sending the 384-well plates to UCLA for use on their qPCR-HRM LightCycler Roche, we need to prepare the plates and reagents separately.
1. Aliquote the amount of DNA into your plate but do not cover! Place in hood overnight to allow the DNA to dry down (buffer will evaporate). Feel free to lightly cover plate with a kim-wipe
2. Thaw Master Mix, water, MgCl2, and primers on ice and keep covered, they are light sensitive
3. Make 2μM primer mix (1μL of the forward primer + 1μL of the reverse primer + 98μL of diH2O)
4. Keep the Primer Mix separate, in it's own clearly labeled tube
5. Make master cocktail to contain the Master Mix, water, MgCl2 and store in its own clearly labeled tube. If the volume is too much for a single tube, keep them in separate tubes but aliquot the amount for the total reaction plus some pipetting error
6. After the DNA is dried, seal the plate with a sticky lid and place in refrigerator until ready to ship
7. Once ready to ship, pack on ice and include reagent tubes wrapped in foil (light sensitive), the DNA plate dried down and sealed, plus the HRM-specific sticky lid
8. Ship OVERNIGHT for next morning delivery to UCLA at the below address
9. When the samples have been sequenced, they will be posted on WebSeq and can be downloaded.

Or general information, you can email the UCLA Human Genetics Core at webseq@genetics.ucla.edu

A general HRM set-up template for keeping notes on the experiment, samples, and master mix.
A general method for analysis of the .ixo files you will retrieve from the WebSeq website.

Suggested analysis method:
The HRM analysis is quick and painless. On the desktop, you will find a shortcut icon to the program LightCycler480. After logging in with the correct name and password, look at the top of the screen to find a drop-down menu. Select "Navigator" and then use the "import" button to select the right .ixo file to import. Following this, look in the left navigation window for the button "Analysis". Of the many options that pop up, select "Tm calling" and hit the "check" button. You then have an option to select the whole plate and press the "calculate" button. Then the melting temperatures (Tm) will be provided. You can select the cells you want and in the bottom left part of the screen, individual Tm's will appear. You can select more information regarding each Tm (e.g. width of peak) but this is essentially your genotype! You need to think about matching up genotypes (presence/absence of specific indels or SNPs) with the denaturing temperature. Good luck and happy genotyping!

2. Quantitative PCR

qPCR assays are used to estimate relative enrichment by abundance of target fragment. This is a relatively inexpensive way to determine if the enrichment (or library prep) was successful prior to sequencing.

Remember that if you are specifically investigating copy number variation in the genome, you need a control gene that is known to be of a single copy.

Here is an example PCR cycle for a generic qPCR:

Program Name

Cycles

Analysis Mode

Target (°C)

Hold (hh:mm:ss)

Ramp Rate (°C/s)

Acquisitions (per °C)

Acquisition Mode

Pre-incubation

1

None

95

00:10:00

4.8

---

None

Amplification

40

Quantification

95

00:00:10

4.8

---

None

60

00:01:00

2.5

---

Single

Melting Curve

1

Melting Curves

95

00:00:10

4.8

---

None

65

00:01:00

2.5

---

None

95

---

---

5

Continuous

Cooling

1

None

40

00:00:10

2

---

None

The PCR cycle program should be applicable to HRM genotyping but as always, the annealing temperature may be altered to match your primer. Another difference is when the acquisition occurs; this is the point in the program at which data (time and temp) is collected. For HRM, this only matters during the final melting curve, whereas in qPCR, this is done repeatedly at the end of every amplification cycle. Check that you are using the appropriate program for your analysis!

In the lab, we use the KAPA kit pretty much as stated in the PDF for quantifying library preps with qPCR.

Sign up for the qPCR run on the LightCycler!

Run all samples in duplicate but the 6 standards in triplicate.

Suggested analysis method:
On the desktop, you will find a shortcut icon to the program LightCycler480. After logging in with the correct name and password, look at the top of the screen to find a drop-down menu. Select "Navigator" and then use the "import" button to select the right .ixo file to import. Next, you will need to import (or set up) your plate template, as to tell the program where your standards are of known concentrations in order to use as a ruler for your own samples. To import your template for qPCR, (for this example we will use Rena's template for quantifying HTS libraries using the KAPA kit), go to the Sample Editor and import the file named kappa-plate-setup-2peat-384-14SAMPLES). Open it up and just scan the headers to check that it looks correct, then select "apply". Following this, look in the left navigation window for the button "Analysis". Of the many options that pop up, select "Absolute quantification/2nd derivative max" in order to do a quantification based on your standards, and hit the "check" button. Make sure to change the efficiency in the menu at the bottom to select the "std curve (in run)" option. Also check that "high sensitivity" is selected in the "high sensitivity" menu. Then click the "Calculate" button. You need to then look at the duplicated you created, first checking the standards and that they are consistent. Make sure to check EVERY standard! Then go ahead and begin surveying your own samples. After all is said and done, you can right-click on the Replicate Statistics Table at the bottom to export.

Also, from here, you can use the concentrations and standards to calculate molarity of your libraries and pool based on that (which is a requirement for all HTS sequencing). Here is an example spreadsheet...but remember, MAKE SURE YOU DISCUSS THIS WITH SOMEONE WHO HAS ALREADY DONE THIS PRIOR TO SUBMISSION TO ANY CORE FACILITY. This is an easy place to go wrong, so double check with your colleagues. Most people will dilute libraries down to 10nM before pooling, but this is flexible, depending on your libraries' concentrations. Best of luck!

Targeted sequencing

If you have enough PCR samples for a 96-well plate, please proceed to the Beckman Coulter Sequencing instructions.

PCR products may be cleaned by gel purification (we use zymoclean kits) or by EXOSAP (as follows).

EXO-SAP
In order to sequence PCR products, excess primers, single stranded DNA, and extra nucleotides (dNTP’s) need to be removed. Exonuclease I (Exo) digests single stranded DNA and Shrimp Alkaline Phosphatase (SAP) dephosphorylates free nucleotides making them unavailable for polymerization. Store these reagents at -20 C.

e. 4 degrees forever
9. Once the BIG DYE reaction is finished, store your cycle sequencing product in the freezer (-20) until ready to send to the CORE.

Sequencing sign up

1. Sign up to sequence at Genoseq CORE.
2. Click on R2R Signup (list) (found on the left hand side) and fill out the required information.
3. Answer YES for request cleaning service!
4. Please bring your reference number with you, remember to check in your samples when you drop them of, and bring back Wayne lab sample racks.
5. Alternatively, you can bring your samples over to the Core and sign up there.

Reduced Representation BS-seq (RRBS) library prep protocol

We have worked through a few protocols and this is showing to be the one that consistently works and may work well with lower input concentrations of DNA. Please see Dr. vonHoldt or Ilana Janowitz before proceeding.
The RRBS protocol can be found here.

1. Remove tubes from incubator, and add 500 µl of buffered phenol. Shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
2. Observe 2 layers. The top layer contains the DNA/RNA and the bottom, organic layer the waste.
3. Remove the top layer and transfer into a new and labeled tube, and discard the bottom layer in the PCI organic waste container.
4. Add 500 µl of PCI to the samples in the new tubes, and shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
5. Observe 2 layers. The top layer contains the DNA/RNA and the bottom, organic layer the waste.
6. Remove the top layer and transfer into a new and labeled tube, and discard the bottom layer in the PCI organic waste container.
7. Add 500 µl of CI to these new tubes, and shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
8. Pipette off the top layer and transfer it to a 1.7ml final epi tube with the final sample name and number.
9. [OPTIONAL] If you want to RNase treat the DNA, add 1ul of a 10 µg/mL stock solution of RNase A to your DNA and incubate at 37ºC for 30-60 min.
10. To this tube add the following:

100 µl of 3M sodium acetate (NaOAc)

1 ml of cold 100% ethanol (EtOH)

11. Invert tubes several times and place in a -20 freezer overnight. (3 hours is sufficient).
12. You can either stop here and do precipitation the next day or after 3 hours, complete the precipitation.

Ethanol Precipitation
1. Remove tubes from the freezer, and centrifuge at max (14,000 rpms for ~10 min) to pellet the DNA.
2. Observe a white or brownish pellet. (May be absent in low concentration samples).
3. Decant the supernatant, being careful not to lose the pellet.
4. Add 1ml of fresh 70% EtOH to the tube containing the pellet. Vortex briefly to re-suspend the pellet.
5. Centrifuge at max (14,000 rpms for 10 min) to pellet the DNA.
6. Decant the EtOH, being careful not to lose the pellet.
7. Remove the remaining 70% EtOH by vacuum centrifuging in the tubes in the Savant Speed-Vac for 10 mins or until the pellet is dry. (Do not use high heat for more than 5 mins).
8. Re-suspend DNA in 100-200 1x TE (or Qiagen’s AE) Buffer.

RNase treatment [if you didn’t do this prior to precipitation]

If you are extracting DNA from an RNAbuffer (e.g. PAXgene RNA tubes which do not contain any DNases), you may consider doing an RNAse treatment of the final DNA*

1. Add 1ul of a 10 µg/mL stock solution of RNase A to your DNA and incubate at 37ºC for 30 min.
2. Recover the DNA by adding 1/10 volume of 3M sodium acetate (pH 6.8) and 2 volumes of isopropanol or 95% ethanol to the DNA containing solution.
3. Incubate on ice for 10 min.
4. Centrifuge at maximum speed for 5 min at room temperature, to pellet the DNA.
5. Discard (carefully) the alcohol. Wash with 70% ethanol and dry DNA via Speed-Vac on low heat for 10 mins.
6. Dissolve in AE, TE or dH2O, as you did in the DNA extraction.

Example: Prepare the diluted ladder for a total of 13 reactions (26μL of 50bp ladder with 234μL water) for the table of trials below

Final ratio of Serapure

Volume of ladder

Volume of Serapure

.5X

20μL

10μL

1X

20μL

20μL

1.2X

20μL

24μL

1.5X

20μL

30μL

1.8X

20μL

36μL

2X

20μL

40μL

2.1X

20μL

42μL

3. Mix gently with a pipette and incubate at room temperature for 5 minutes.
4. Place tubes on magnetic stand and incubate for 5 minutes. Beads should be drawn out of solution and should gather on the side of the tube in contact with the magnet.
5. Carefully pipette off supernatant - without disturbing the beads - and discard.
6. Add 500uL of fresh 70% ethanol to each tube, incubate for 1 min, pipette off supernatant and discard.
7. Repeat step 6.
8. Allow beads to air dry for 10 min with tube caps open to remove all traces of ethanol or place beads of a 37°C heat block for 3-4 min until dry.
9. Resuspend beads in 20μL EB buffer to elute DNA (tubes are not on the magnet for this step). Vortex/flick the tubes vigorously to ensure proper mixing of beads with DNA
10. Incubate on magnetic stand for 5 min.
11. Transfer supernatant to newly labeled tubes and discard beads.
12. Mix supernatant with 3μL of loading dye.
13. Electrophorese in a 1.5% agarose gel for 60min at 100V (use the same ladder (3μL with 2μL 6X dye) in the gel).
14. Compare the results of the trial volumes do obtain the most appropriate XμL needed to do the following AMPure Bead Cleanup Protocol or for library preps.

AMPure XP bead clean-up of DNA

1. Use sample DNA concentration to calculate volume that will yield 1μg of DNA.

To calculate the number of microliters needed to obtain 1μg of DNA, divide 1000 by the concentration of DNA in ng/μL.

To obtain genomic DNA, use a ratio of XuL bead solution per μg of sample DNA (X is the optimum volume figured out by testing the ampure mixture; typically 40-60μL).

2. Obtain clear 1.7mL tubes. In each 1.7mL tube, combine bead solution, DNA, and 1X TE buffer in the following ratio: XμL : 1μg : up to 160μL total volume.

Can be scaled up to combine as much as 3μg DNA with 3XμL bead solution and 1X TE to a total volume of 480μL in each tube.

3. Mix gently with a pipette and incubate at room temperature for 5 minutes.
4. Place tubes in magnetic tube holder and allow to rest for 2 minutes. Beads should be drawn out of solution and should gather on the side of the tube in contact with the magnet.
5. Carefully pipette off supernatant - without disturbing the beads - and discard.
6. Add 500μL fresh 70% ethanol to each tube, then pipette off ethanol and discard.
7. Repeat step 6.
8. Allow beads to air dry with tube caps open to remove all traces of ethanol (~5-10 minutes) or place beads of a 37°C heat block for 3-4 min until dry. You can also use a sterile toothpick to remove any blobs of ethanol from the tube walls.
9. Resuspend beads in 10μL 1X TE buffer to elute DNA.
10. Transfer elute to new labeled tubes (combining duplicates from the same initial sample, if possible) and discard beads.
11. NanoDrop to determine new sample concentration. Perform ethanol precipitation to concentrate if necessary.

RAD-seq

Please print out a RAD-seq protocol for your own record keeping. Please check with others who have experience with this protocol before conducting any large-scale library prep effort. There are many nuances and sticky spots that should you should be aware of prior to beginning lab work.

RRBS

Please print out a RRBS protocol for your own record keeping. Please check with others who have experience with this protocol before conducting any large-scale library prep effort. There are many nuances and sticky spots that should you should be aware of prior to beginning lab work.